ION TRANSPORT IN CHLOROPLASTS WITH
ROLE IN REGULATION OF PHOTOSYNTHESIS
ANDREI HERDEAN
FACULTY OF SCIENCE
DEPARTMENT OF BIOLOGICAL AND ENVIRONMENTAL SCIENCES
Akademisk avhandling för filosofie doktorsexamen i Naturvetenskap med inriktning Biologi,
som med tillstånd från Naturvetenskapliga fakulteten kommer att offentligt försvaras
fredagen den 2 oktober kl. 10.00 i Hörsalen, Institutionen för biologi och miljövetenskap, Carl
Skottsbergs gata 22B, Göteborg.
Examinator: Professor Adrian Clarke, Institutionen för biologi och miljövetenskap, Göteborgs
Universitet
Fakultetsopponent: Professor Giovanni Finazzi, Laboratoire de Physiologie Cellulaire
Végétale, Université de Grenoble, France
ISBN: 978-91-85529-85-8
© Andrei Herdean, 2015
ISBN: 978-91-85529-85-8
Printed by Kompendiet, Göteborg 2015
For my family!
“I may not have gone where I intended to go, but I think I have
ended up where I needed to be.”
Dirk Gently
Ion transport in chloroplasts with role in regulation
of photosynthesis
Andrei Herdean
University of Gothenburg, Department of Biological and Environmental Sciences
Box 461, SE-40530 Gothenburg, Sweden
ISBN:978-91-85529-85-8
ABSTRACT
Photosynthesis is the primary energy source of almost all ecosystems on
Earth. Oxygenic photosynthesis has appeared approximately 2.4 billion years ago
and has since then gradually evolved into the complex process that land plants,
algae and cyanobacteria perform today. During the course of evolution, these
organisms have also developed mechanisms to improve photosynthetic efficiency, to
cope with changes in the environment, and effectively use the available resources.
The environment in which photosynthetic organisms grow contains numerous
ionic compounds. These compounds are taken up and used in numerous important
processes, including photosynthesis in chloroplasts. As in the rest of the cell,
specialized proteins named channels and transporters mediate ion transport across
membranes and control ion homeostasis in chloroplasts.
The work presented in this thesis addresses the role in regulation of
photosynthesis of ion channels and transporters from the chloroplast inner envelope
(Paper I), and the thylakoid membrane (Paper I to V) of Arabidopsis thaliana.
Potassium ion fluxes mediated by the chloroplast K+/H+ antiporters KEA1, KEA2 and
KEA3 regulate the composition of the proton motive force (PMF) across the thylakoid
membrane that activates photoprotective mechanisms (NPQ) (Paper I). In Paper II,
an Arabidopsis mutant named pam71 is found disturbed in photosystem II efficiency
and the adjustment of PMF, due to altered Ca2+ homeostasis in the chloroplast. In
Paper III, it is shown that the thylakoid phosphate transporter PHT4;1 affects the
availability of phosphate for ATP synthesis, and also alters NPQ activation kinetics
and PMF composition. A novel thylakoid voltage-dependent chloride channel
(VCCN1) is identified in Paper IV, and shown to affect PMF and NPQ activation
upon illumination and after rapid shifts from low light to high light. In Paper V it is
shown that the thylakoid chloride channel CLCe contributes to the modulation of
PMF as well as to the regulation of electron transfer and state transition.
Taken together, the findings of this thesis bring novel mechanisms of anion
and cation transport across the thylakoid membrane and the chloroplast inner
envelope with role in regulation of photosynthesis.
LIST OF PUBLICATIONS
This thesis is based on the following papers, which are referred to by their Roman
numerals in the text:
I.
II.
III.
IV.
V.
Kunz HH, Gierth M*, Herdean A*, Satoh-Cruz M, Kramer DM, Spetea C &
Schroeder JI (2014) Plastidial transporters KEA1,-2, and-3 are essential for
chloroplast osmoregulation, integrity, and pH regulation in Arabidopsis.
Proceedings of the National Academy of Sciences USA 111: 7480-7485.
Schneider A, Herdean A, Steinberger I, Morper A, Labs M, Spetea C & Leister
D. The photosynthesis-affected mutant71 of Arabidopsis influences
photosystem II efficiency and the proton motive force composition.
Manuscript.
Karlsson PM*, Herdean A*, Adolfsson L, Beebo A, Nziengui H, Irigoyen S,
Űnnep R, Zsiros O, Nagy G, Garab G, Aronsson H, Versaw WK & Spetea C
(2015). The Arabidopsis thylakoid transporter PHT4;1 influences phosphate
availability for ATP synthesis and plant growth. The Plant Journal, doi:
10.1111/tpj.12962**.
Herdean A, Teardo E, Nilsson AK, Pfeil BE, Johansson ON, Űnnep R, Zsiros
O, Nagy G, Dana S, Solymosi K, Garab G, Szabó I, Spetea C* & Lundin B*. A
voltage-dependent chloride channel fine-tunes photosynthesis in plants.
Manuscript.
Herdean A, Nziengui H, Zsiros O, Garab G, Lundin B & Spetea C. The
Arabidopsis thylakoid chloride channel AtCLCe contributes to chloride
homeostasis and photosynthetic regulation. Manuscript.
Other papers not included in this thesis:
Yin L, Fristedt R, Herdean A, Solymosi K, Bertrand M, Andersson MX,
Mamedov F, Vener AV, Schoefs B & Spetea C (2012) Photosystem II function
and dynamics in three widely used Arabidopsis thaliana accessions. PloS
One 7: e46206-e46206.
Flood PJ, Yin L, Herdean A, Harbinson J, Aarts MG, & Spetea C (2014)
Natural variation in phosphorylation of photosystem II proteins in Arabidopsis
thaliana: is it caused by genetic variation in the STN kinases? Philosophical
Transactions of the Royal Society B: Biological Sciences 369: 20130499.
Fristedt R, Herdean A, Blaby-Haas CE, Mamedov F, Merchant SS, Last RL &
Lundin B (2015). PHOTOSYSTEM II PROTEIN33, a protein conserved in the
plastid lineage, is associated with the chloroplast thylakoid membrane and
provides stability to photosystem II supercomplexes in Arabidopsis. Plant
Physiology 167: 481-492.
* Shared authorship
** Reprinted with permission from John Wiley and Sons
LIST OF ABBREVIATIONS
AGI
CBB
CLC
Cyt
DCMU
DIRK
ECS
ETC
ETR
e−
F0
Fm
gH+
H+
KEA
LHC
NADPH
NADP+
NPQ
OEC
pam
PC
PHT
Pi
PMF
PS
WOC
VCCN
WT
 (II)
pH

H+
Arabidopsis gene index
Calvin Benson Bassham
Chloride channel
Cytochrome
3-(3,4-dichlorophenyl)-1,1-dimethylurea
Dark interval relaxation kinetics
Electrochromic band shift
Electron transfer chain
Electron transfer rate
Electrons
Minimum fluorescence level at time 0
Maximum fluorescence level after a saturating pulse
H+ conductivity of thylakoid ATP synthase
Protons
Potassium efflux antiporter
Light-harvesting complex
Reduced nicotinamide adenine dinucleotide phosphate
Oxidized nicotinamide adenine dinucleotide phosphate
Non-photochemical quenching
Oxygen-evolving complex
Photosynthesis-affected mutant
Plastocyanin
Phosphate transporter
Inorganic phosphate
Proton motive force
Photosystem
Water-oxidizing complex
Voltage-dependent chloride channel
Wild type
PSII quantum yield
Difference in pH between thylakoid lumen and stroma
Difference in electrical potential between thylakoid lumen and stroma
Steady-state proton flux via ATP synthase
TABLE OF CONTENTS
Page
1. Introduction
1.1.
Photosynthesis
1.2.
Chloroplast organization
1.3.
Light-harvesting
1.4.
Photosystem II
1.5.
Cytochrome b6f
1.6.
Photosystem I
1.7
ATP synthase
1.8.
Regulation of photosynthesis
1.9.
Classification of ion transporters
1.9.1. Channels and porins
1.9.2. Secondary transporters
1.9.3. Primary transporters
1
2
3
4
5
5
5
6
7
7
7
9
2. Scientific aims
13
3. Methodology
3.1.
Arabidopsis thaliana as a model plant
3.2.
Functional genomics as approach to study transporters
3.3.
Chlorophyll fluorescence as approach to study
photosynthesis
3.3.1
Fast kinetics of fluorescence induction
3.3.2
Pulse amplitude modulated fluorescence
3.4.
Electrochromic band shift
17
17
19
19
20
22
4. Role of ions and ion transporters in photosynthesis
4.1.
Ion transport with role in regulation of the proton motive
force
4.1.1
Potassium
4.1.2
Calcium
4.1.3
Phosphate
4.1.4
Chloride
4.1.5
Other ions
4.2.
Regulation of electron transfer
4.3.
Regulation of thylakoid architecture
4.4.
Role in signalling and enzymatic activation
4.5.
Regulation of photoprotective mechanisms
29
30
30
33
34
34
34
35
36
5. Conclusions and outlook
39
6. Acknowledgements
43
7. References
47
29
INTRODUCTION
1
Ion transport in chloroplasts with role in regulation of photosynthesis
1.1 Photosynthesis is the process by which solar energy is absorbed by some living
organisms (plants, algae and certain bacteria), converted and stored as chemical
energy.
Based on geological evidence photosynthesis-capable organisms are estimated
to have appeared more than 3 billion years ago. The first organisms to successfully
utilize energy from sunlight were most likely anoxygenic (non-oxygen evolving), and
used hydrogen peroxide as the primary electron donor, and before that ferrous iron
[1]. The greatest leap in innovation of the photosynthetic mechanism was the
transition to oxygenic photosynthesis. The use of water as the primary electron
source, and the subsequent release of O2 changed the composition of the Earth’s
atmosphere starting approximately 2.4 billion years ago [2]. Oxygenic photosynthesis
and its by-product O2 changed Earth’s environment to such an extent that it affected
the evolution of life, allowing for organisms to grow in size and complexity (Fig. 1).
During millennia photosynthesis evolved and diversified to become an efficient
energy-harvesting mechanism in all environments.
Figure 1. Size of fossils throughout history and the two sharp increases in
atmospheric O2 from photosynthesis. Oxygen is shown as percentage of present
atmospheric levels (PAL) during the Archaean, Proterozoic and Phanerozoic
geological eons. Adapted from ”Two-phase increase in the maximum size of life over
3.5 billion years reflects biological innovation and environmental opportunity.” by JL
Payne et al. (2009) Proc Natl Acad Sci USA 106: 24-27. Adapted with permission.
Oxygenic photosynthesis is a complex process where many simultaneous
mechanisms interact with each other to culminate with the reduction of inorganic
carbon, which in turn is converted into organic molecules to build cellular
components. Part of the photosynthetic machinery has evolved mechanisms for
1
Andrei Herdean
efficient light harvesting, oxidation of water, reduction of NADP+ and phosphorylation
of ADP; this sum of processes is known as the light-dependent reactions. Another
part of the photosynthetic machinery uses ATP and NADPH, to fix CO2 and
synthesize a 3-carbon sugar (triose phosphate), during the so-called lightindependent reactions or the Calvin-Benson-Bassham (CBB) cycle. The work
presented in this thesis will focus mostly on the machinery that performs the light
reactions (Paper I to V), with some emphasis on the CBB cycle (Paper II and III).
1.2 Photosynthesis in land plants and algae takes place in a specialized organelle
called chloroplast. Chloroplasts are plastids that are thought to be the result of an
endosymbiotic event between an early eukaryote and cyanobacteria. This
endosymbiosis is considered to have taken place after the one yielding mitochondria
since all eukaryotes have mitochondria, but not all have chloroplasts.
Two distinct membranes separate the chloroplast stroma from the cytosol: the
outer envelope membrane and the inner envelope membrane. The outer envelope is
a highly permeable membrane that originated from the eukaryotic host, whereas the
inner envelope is less permeable but with a high number of transport proteins, and
has originated from the ancestral cyanobacteria. The third membrane system in
chloroplasts is the photosynthetic membrane, also called the thylakoid membrane.
The thylakoid membrane separates the two solute compartments of the chloroplasts
the acidic thylakoid lumen, and the alkaline chloroplast stroma.
A specific organization of the thylakoid membrane (Fig. 2), also called
thylakoid ultrastructure, exists in order to maximize light harvesting, energy transfer
and repair mechanisms [3]. Thylakoids have two distinct morphological and
functional regions called grana and stroma lamellae. The thylakoid membrane
contains all the protein complexes performing the light-dependent reactions
photosystem II (PSII), cytochrome b6f (Cyt b6f), photosystem I (PSI), and ATP
synthase. These protein complexes are distributed between the grana and stroma
lamellae, with PSII found predominantly in the grana region and Cy t b6f, PSI and
ATP synthase found predominantly in the stroma lamellae (Fig. 2). PSII and PSI are
additionally connected to distinct light harvesting protein-pigment complexes, which
in turn follow the distribution of PSII and PSI throughout the membrane. Despite that
PSII and PSI are spatially separated in thylakoids, in certain situations these two
complexes together with their light harvesting antennas can come in close proximity
and transfer excitation energy from one complex to the other in a process called spillover [4]. Furthermore, another similar mechanism that implies changes in thylakoid
ultrastructure is state transition [5], a process regulated by phosphorylation of light
harvesting antennas as well as anion distribution between chloroplast stroma and
thylakoid lumen (Paper V).
The chloroplast stroma is the site of the light-independent reactions (CBB
cycle), which despite its name occurs only during the day since they consume ATP
and NAPDH generated by the light-dependent reactions. The CBB cycle has 3
2
Ion transport in chloroplasts with role in regulation of photosynthesis
phases: carbon fixation, reduction reactions, and ribulose 1,5-biphosphate
regeneration.
Figure 2. Schematic representation of the chloroplast ultrastructure. Figure
presents the chloroplast photosynthetic membrane (thylakoid membrane) organized
into grana and stroma lamellae. Distribution of photosynthetic protein complexes
between grana stacks (appressed thylakoids) and stroma lamellae (non-appressed
thylakoids) is shown, with photosystem II (PSII) found in grana regions and PSI,
cytochrome b6f (Cyt b6f) and ATP synthase in the stroma lamellae, and to a lesser
extent in grana stacks. Chloroplasts have two soluble compartments: thylakoid lumen
(grey) and chloroplast stroma (white). In the chloroplast stroma carbon fixation takes
place via the CBB cycle, shown with circular black arrows.
1.3 Light-harvesting antennas capture the energy carried by photons using a
special set of photosynthetic pigments. There are three basic classes of pigments
used in light harvesting: chlorophylls, carotenoids and phycobilins. Chlorophylls are
considered to be the most important, with chlorophyll a the most significant one.
Chlorophylls are essential because they form a porphyrin ring where electrons (e−)
are free to move. When an e− in a chlorophyll a porphyrin ring receives excitation
energy from the antenna, its energy level becomes elevated, and the e− moves to a
different molecule towards the terminal electron acceptor. Carotenoids are usually
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Andrei Herdean
red, orange or yellow and are composed of two small six-carbon rings connected by
a chain of carbon atoms. Carotenoids transfer excitation energy to chlorophylls and
not directly to the electron transfer chain. Phycobilins are only found in
Cyanobacteria and Rhodophyta, and unlike the other pigments, these are soluble
and not present in membranes. Instead, they form mobile light-harvesting antennas
known as phycobilisomes.
Antennas are highly regulated and very diverse among different organisms. By
increasing the size of the light-harvesting antenna, an organism is able to increase
the absorption cross-section for light without having to build more reaction centers,
which would be costly in terms of cellular resources. Additionally, by varying the
pigment composition more of the light spectrum can be trapped. Connectivity
between antennas (energy collector) and reaction centers (energy sink) is a common
regulatory mechanism that organisms use to maximize harvesting efficiency or
protect against photo-damage.
Plants have two major light-harvesting complexes, each associated with either
PSII (LHCII) or PSI (LHCI). LHCII but not LHCI has been shown to be able to
disconnect from PSII and connect to PSI during state transitions [5], hence
increasing the PSI antenna size. This process is reversible and is triggered in
response to changes in light quality. The dynamics of LHCII movement is regulated
by phosphorylation, and subsequently by changes in thylakoid architecture, as well
as ion fluxes (Paper V).
1.4 Photosystem II (PSII) is a protein supercomplex that exists as a dimer in vivo
and can have in excess of 30 subunits, depending on the organism [6]. It is located in
the thylakoid membrane and found predominantly in the grana stacks in land plants
(Fig. 2). The core of PSII, defined as the minimum number of proteins required to
oxidize water, is highly conserved across all photosynthetic organisms, and it
consists of D1, D2, CP43, CP47 and cyt b559 subunits [6] (Fig. 3). The oxygenevolving complex (OEC) or water-oxidizing complex (WOC) is the Mn4CaO5
inorganic cluster located at the donor side of PSII that splits 2 H2O molecules into O2,
4 H+ and 4 e-. The OEC is the primary electron source for the electron transfer chain
(ETC). Since its discovery [7-9], the OEC has been subject for many studies to solve
the exact mechanism by which it oxidizes water. The original model proposed by Kok
and co-workers [8, 9] is still used today and assumes four intermediate steps (S
states: S0 to S4), i.e., four conformational changes of the inorganic cluster that result
in the release of O2. The exact position of the Mn4CaO5 atoms in each of the S states
as well as its assembly remain unclear today, despite the extensive work to solve it
[10-12].
Electrons from the OEC are shuttled to tyrosine Z (Yz), the primary electron
acceptor, an amino acid that is part of the D1 subunit of PSII. From Yz electrons
move to a chlorophyll molecule named P680. P680 channels excitation energy from
the light-harvesting antennas and elevates the energy levels of the electron before it
moves to pheophytin (Phe), which is another chlorophyll molecule but without the
4
Ion transport in chloroplasts with role in regulation of photosynthesis
central Mg2+. Further the electrons move to a quionone molecule QA, and then to QB
before transfer to the Cyt b6f complex by a mobile plastoquinone (PQ) molecule. PQ
takes 2 e− from PSII and 2 H+ from the chloroplast stroma, by this adding 2 more H+
to the trans-thylakoid proton gradient.
PSII is often the primary target of photodamage, and is also the complex with
the highest turnover rate in chloroplasts even in normal growth conditions. PSII
function and dynamics are additionally altered when the thylakoid ionic environment
is changed. Oversaturation of the chloroplast with K+ reduces PSII activity (Paper I),
and excess of Ca2+ in the thylakoid lumen can cause instability and low activity of the
OEC (Paper II). Changes in chloroplast anion (Cl-) homeostasis impacts much less
PSII function (Paper IV, V).
1.5 Cytochrome b6f is the protein complex that links PSI and PSII by oxidizing the
lipophilic plastoquinol and reducing the soluble plastocyanin (Fig. 3). It functions as a
dimer and can have 8 to 9 subunits depending on the organism [13, 14]. Cyt b6f
mediates the linear and cyclic electron transport that is coupled with H+ translocation,
significantly contributing to the transmembrane H+ gradient required for ADP
phosphorylation. Cyclic electron transport does not generate NADPH, and it involves
oxidation of PSI-reduced ferredoxin (Fd) [15] or ferredoxin:NADP+ oxidoreductase
(FNR) [16].
1.6 Photosystem I, unlike PSII, is predominantly located in the non-appressed
regions of the thylakoid membrane (Fig. 2), and it catalyses a light-dependent
electron transport from plastocyanin to ferredoxin (Fig. 3). It consists of up to 14
subunits [17], and has been found in monomeric as well as trimeric form [18, 19].
Light-dependent charge separation occurs in a chlorophyll a molecule named P700,
which channels excitation energy from the PSI-specific light-harvesting antenna
LHCI. Additional light-harvesting antennas (LHCII) can in certain conditions connect
to the PSI antenna complexes in response to changes in light quality or intensity, in a
process called state transition [20]. Electron transport within PSI starts with P700 that
is considered to be the primary electron donor, which in turn takes its electrons from
PC. Excited P700 transfers an electron to another chlorophyll molecule termed A0,
followed by a phylloquinone termed A1, and then to 3 iron-sulphur centres FX, FA and
FB before reaching the PSI terminal electron acceptor ferredoxin. Ferredoxin
ultimately transfers electrons to FNR, which will reduce NADP+ to NADPH that is
used in the CBB cycle. In contrast to P680 which is considered the strongest
biological oxidizing agent, P700 is considered the strongest biological reducing
agent.
1.7 ATP synthase is a large protein complex consisting of two distinct parts: the
catalytic hydrophilic CF1, and the transmembrane H+-translocating CF0. CF1
consists of 9 proteins separated in 5 groups (3, 3, 1, 1 and 1) [21], whereas
5
Andrei Herdean
CF0 consists of 4 distinct proteins (I, II, III and IV), where protein III is found in 12-14
copies [22, 23]. The ATP synthase is a rotary type of protein that uses the
electrochemical H+ gradient generated by the PSII-Cytb6f-PSI electron transfer chain
in order to ultimately phosphorylate ADP into ATP (Fig. 3). The CF0 subunit IV
channels H+ from the thylakoid lumen to the chloroplast stroma and uses the
resulting mechanical energy to generate ATP, which is subsequently used in the
CBB cycle.
Ion transport can have a direct influence on ATP synthase activity by either
limiting the supply of inorganic phosphate (Pi) to the catalytic site (Paper III), or by
changing the composition of the proton motive force (Paper IV, V), which drives ATP
synthesis.
Working together PSII, Cyt b6f, PSI and the ATP synthase orchestrate an
elegant mechanism that splits H2O molecules using energy collected from photons,
to finally generate ATP and NADPH.
Figure 3. Schematic representation of the electron transfer chain. Small black
arrows indicate the linear electron transfer chain starting with the OEC at PSII and
ending with FNR at PSI. Red arrows indicate the Q cycle. Grey arrows in PSI
indicate an alternative electron transfer pathway. Black dashed arrows indicate H+dependent ADP phosphorylation. Grey dashed arrows indicate reversible ATP dephosphorylation.
1.8 Regulation of photosynthesis on a metabolic level is performed by the
circadian clock. Chloroplast gene expression and CO2 fixation have been shown to
be tightly influenced by plant circadian rhythms [24]. The chloroplast has, however,
developed additional specialized regulatory mechanisms for fine tuning the
photosynthetic efficiency and for response to stress.
Light intensity and composition in natural environments changes quickly and
to a large extent during the day, and because of this, plants have developed
6
Ion transport in chloroplasts with role in regulation of photosynthesis
photoprotective mechanisms to avoid photodamage, or photoinhibition [25]. Under
excess light conditions the primary target of photoinhibition is PSII, whereas under
certain conditions (low temperature and low light) PSI is more prone to damage [26].
In response to fast fluctuations in light intensity, photosynthetic organisms have
developed similarly fast responses in order to avoid excess excitation energy being
channelled from light-harvesting antennas to the reaction centers.
In a relaxed, non-stressful state, light-harvesting antennas are connected to
the reaction centers (RCs). When a fast increase in light intensity takes place
antennas get disconnected, and by doing this, the RCs become protected from
excess excitation energy. The process of antenna quenching is controlled by direct
protonation [27] or phosphorylation [5] of LHC, protonation of PsbS [28, 29], and
violaxanthin de-epoxidation [30]. Because the thylakoid membrane is extremely
protein dense, mobility of LHC is restricted, and hence dynamic and reversible
changes in thylakoid membrane architecture occur [31], concomitantly with antenna
quenching.
Acidification of the thylakoid lumen is the trigger for most of the
photoprotective responses. Ion transporters in turn regulate fast dynamic
adjustments of the lumenal pH by import and export of K+ [32, 33], Paper I), and by
regulating the Pi availability to ATP synthase (Paper III).
Long-term acclimation of photosynthetic efficiency is another regulatory
mechanism, and it involves adjustments of pigment composition and content. Under
low light conditions, plants make more chlorophyll to enhance light harvesting and
under extended excess light, they reduce chlorophyll content.
1.9 Ion transport proteins belong to three categories with several subcategories in
the Transport Classification DataBase (http://www.tcdb.org/, [34]).
1.9.1 Channels and porins constitute the non-energy consuming transport system,
and carry solutes down their electrochemical gradient. This class of proteins
transport molecules at a very fast rate of 107 - 108 molecules s-1. Regulation of
opening and closing of -type channels varies depending on the subcellular location
and the type of substrate. In chloroplasts, Ca2+- (TPK3 in [33]) and voltage- (VCCN1
in Paper IV) gating are common ways of regulating channel activity. CLCe belongs to
the chloride channel CLC family ([35, 36] and Paper V), but the formal proof of its
channel activity as well as its regulatory mechanism is still missing. The β-barrel
porins described in the chloroplast to date include exclusively outer envelope
proteins responsible for import of amino acids, primary amines, ATP and inorganic
pyrophosphate [37].
1.9.2 Secondary transporters require the gradient of a co-transported solute for
transport of the compound of interest. This class of proteins include antiporters
(transport of two solutes in opposite directions) and symporters (transport of two
7
Andrei Herdean
solutes in the same direction), and work at a slower rate than channels, 102 - 104
molecules s-1. Both antiporters and symporters transport down the electrochemical
gradient one type of solute and another type against its electrochemical gradient. In
chloroplasts, this type of mechanism is used for transport of phosphate (e.g., PHT4;1
in Paper III), potassium (e.g., KEAs in Paper I), sodium, amino acids and ATP / ADP
exchange.
Figure 4. Schematic map of ion transporters from the chloroplast. Transporters
located in the thylakoid membrane are found on the stroma lamellae region of
thylakoids rather than in the stacked grana (not represented in the figure). The
chloroplast outer envelope does not have ion transporters, being easily permeable
for most solutes. Adapted from ”Ions channels/transporters and chloroplast
regulation.” by G. Finazzi et al. (2014) Cell Calcium 58: 86-97. Adapted with
permission.
8
Ion transport in chloroplasts with role in regulation of photosynthesis
1.9.3 Primary transporters consume energy in the form of ATP or electrochemical
gradient to accomplish transport across membranes. This type of transporters is the
slowest one with a rate of 10 - 103 molecules s-1. Photosynthetic reaction centers,
PSII and PSI, are considered in this category due to their role in translocation of eacross the thylakoid membrane. The ATP synthase is another complex defined as a
primary transporter because of its H+-translocating properties. Cu2+ and Zn2+ are also
transported via ATPases that consume ATP to drive transport against their
concentration gradient.
As in the rest of the cell, due to their relative impermeable membranes,
chloroplasts have highly specialized transport proteins (Fig. 4, Table 1) for transport
of ions across the inner envelope and thylakoid membrane. To date 28 such proteins
have been found in Arabidopsis thaliana (reviewed in [38]), and an additional channel
is identified in this thesis (Paper IV). A Ca2+ efflux activity is also found in the
thylakoid membrane (Paper II), and it remains to be elucidated which is the protein
responsible and to which transport category it belongs.
Protein
classification
Channels
Secondary
transporters
Primary
transporters
TPK3
At4g19160
Chloroplast
location
Thylakoid
CLCe
At4g35440
Thylakoid
Cl-, NO3-?
VCCN1
MSL2
MSL3
GLR3.4
MSR2-11
#
At5g10490
At1g58200
At1g05200
At5g22830
TAAC/ PAPST1
At5g01500
Cl-, NO3Ca2+, Na+, H+
Ca2+, Na+, H+
Ca2+
2+
Mg
ATP/ADP;
PAPS
K+
Protein name
AGI number
Substrate
+
K
KEA3
At4g04850
PHT4;1
At2g29650
KEA1
At1g01790
Thylakoid
Envelope
Envelope
Envelope
Envelope
Thylakoid/
envelope
Thylakoid
Thylakoid/
envelope
Envelope
KEA2
At4g00630
Envelope
K+
PHT2;1
PHT4;4
PHT4;5
Cs Nitr1-L
SULTR 3;1
MAR1
NAP14
YLS4
YLS6
PIC1
NiCo
NHD1
CF0
HMA8
At3g26570
At4g00370
At5g20380
At1g68570
At3g51895
At5g26820
At5g14100
At5g41000
At3g27020
At2g15290
At4g35080
At3g19490
AtCg00130
At5g21930
Envelope
Envelope
Envelope?
Envelope
Envelope
Envelope
Envelope
Envelope
Envelope
Envelope
Envelope
Envelope
Thylakoid
Thylakoid
HMA1
At4g37270
Envelope
HMA6
ACA1
At4g33520
At1g27770
Envelope
Envelope
HPO42HPO42-, ASC
2HPO4
NO3
2SO4
2+
Fe
2+
Fe
Fe2+
2+
Fe
Fe2+
2+
Fe
Na+
H+
Cu2+
2+
2+
Cu , Zn ,
2+
Ca
Cu2+
Ca2+
HPO42K+
Reference
[33]
[35, 36]
Paper V
Paper IV
[39]
[39]
[40]
[41]
[42-44]
[32, 45], Paper I
[46-49],
Paper III
[45], Paper I
[45, 50],
Paper I
[51, 52]
[53, 54]
[53]
[55]
[56]
[57]
[58]
[59, 60]
[59, 60]
[61, 62]
[62]
[63]
[64, 65]
[48, 66]
[67]
[66, 68]
[69]
Table 1. List of known transporters from the chloroplast. The above list excludes
the photosynthetic reaction centers PSII and PSI, due to their principal role in etransport rather than ions. # AGI number is only given in Paper IV.
9
SCIENTIFIC AIMS
2
Ion transport in chloroplasts with role in regulation of photosynthesis
The aim of the work presented in this thesis is to investigate the role of chloroplast
ion transport in the context of photosynthesis.
Specific aims:
Paper I
To establish the physiological role in Arabidopsis of the envelopelocated K+/H+ antiporters KEA1 and KEA2, and the thylakoid-located
KEA3 for photosynthesis and plant growth.
Paper II
To study the phenotype of the Arabidopsis photosynthesis-affected
mutant71 (pam71), in relation to Ca2+ homeostasis in the chloroplast.
Paper III
To investigate the importance in Arabidopsis of the thylakoid-located
phosphate transporter PHT4;1 for ATP synthesis and plant growth.
Paper IV
To characterize the role in photosynthesis of a newly identified voltagedependent chloride channel (VCCN1) localized to the Arabidopsis
thylakoid membrane
Paper V
To further investigate the role of the thylakoid-located chloride channel
CLCe in photosynthesis.
13
METHODOLOGY
3
Ion transport in chloroplasts with role in regulation of photosynthesis
3.1 Arabidopsis thaliana as a model plant. Arabidopsis thaliana is a small
flowering plant native to Eurasia, and it is part of the Brassicaceae family. The first
scientific publication about this plant dates back to 1907 and was published by
Laibach [70], who characterized its chromosomes. Later on (1943) he published
another article about Arabidopsis, pointing out the advantages of working with this
plant [71]. In 1965 the first Arabidopsis conference was held, and in 1986 the first
transformation with Agrobacterium tumefaciens was reported [72] as well as the first
gene sequence [73]. In 2000 the complete sequence of its genome was published
[74], and the potential of T-DNA insertion knockout mutants was demonstrated as
early as 1999 [75]. Several institutions have in the past years created T-DNA
insertion populations with the aim of providing knockout mutants for all of its 27,000
genes.
Some of the most important advantages of working with Arabidopsis thaliana
are the following: relatively small genome (135 Mbp); the first plant genome
completely sequenced; small size of the plant and short life cycle, making it suitable
for transformation with Agrobacterium tumefaciens; and the availability of an
extensive literature of more than 100 years of research.
3.2 Functional genomics as approach to study transporters. Due to current
advances in whole genome sequencing and the fast development of the growing field
of bioinformatics, identification of putative genes coding for transporters or channels
has become a matter of data mining. In order to identify and characterize new
chloroplast-localized transporters, the first step is to check for sequence similarity
with known transporters against a database of genes that code for predicted
membrane proteins (e.g., http://aramemnon.uni-koeln.de/); it is unlikely for a
transporter or channel not to have any transmembrane regions. Secondly, an
analysis of the transit peptide sequence will indicate the probability of the chosen
sequence to code for a protein localized to the chloroplast. Transit peptides act as
targeting sequences that guide the newly-synthesized protein to the appropriate
organelle. These sequences vary greatly in length, composition and organization
[76]. However, there are readily available algorithms that analyse and predict the
destination of the transit peptide (e.g., TargetP, ChloroP, iPSort, WoLF-PSort,
MultiLoc).
Once a suitable candidate gene is identified, its subcellular localization needs
to be confirmed by a microscope approach using green fluorescence protein (GFP)
transformed plants, and western blot (WB) using GFP- or candidate protein specific
antibody. Confirmation of correct localization of the protein is done by isolating pure
chloroplast inner envelope membranes, as well as thylakoid membranes against
which the specific antibodies are used in WB analysis (Fig. 5). In the absence of a
candidate protein specific antibody, the expression profile of the candidate gene
(method used in Paper IV) needs to be analysed to exclude the possibility of the
gene not being expressed in chloroplasts but in other plastids in different parts of the
plants (e.g., flowers, roots etc.).
17
Andrei Herdean
Establishing the transporter function is of crucial importance in order to further
investigate the physiological role of the protein. Heterologous expression of the
native gene and electrophysiological characterization can confirm the substrate
specificity and properties of the newly identified ion transporter (method used in
Paper IV).
Furthermore, a reverse genetics approach is taken to validate the
physiological role (method used in Papers I to V). Unlike forward genetics that is
used to answer the question “What is the genotype of the observed altered
phenotype?”, reverse genetics aims at answering the question “What is the
phenotype of a specifically altered gene?”. Because of the currently available
bioinformatics tools and the extensive T-DNA insertion libraries, reverse genetics is a
very efficient approach to study uncharacterized genes. Knowing the substrate of a
novel ion transporter protein, as well as its subcellular location, specific experiments
can be designed to investigate the role of the protein in chloroplast metabolism and
photosynthesis.
Figure 5. Schematic representation of in silico and experimental approaches
used in functional genomics.
An alternative approach to identification of candidate genes from genomic
databases is screening of chloroplast proteomics databases. Proteomics has been
proven as a successful approach for identification of a large number of chloroplast
proteins [48, 77-81]. Nevertheless, due to low abundance of transporter and channel
proteins, especially in the thylakoid membrane, where relative to photosynthetic
proteins, they are predicted to be found in 107-1014 fold lower quantities, proteomics
approaches may not always be sensitive enough to detect such proteins.
18
Ion transport in chloroplasts with role in regulation of photosynthesis
3.3 Chlorophyll fluorescence as approach to study photosynthesis. Chlorophyll
a fluorescence, also known as the Kautsky effect, was discovered in 1931 by Hans
Kautsky, who observed chlorophyll a fluorescence coming from leafs with the naked
eye [82]. Based on his observations, he later proposed that there are two reactions
taking place during illumination, presumably charge separation in PSII and PSI [83,
84]. The fluorescence signal is now thought to arise from an energetic back-reaction
of the PSII core chlorophyll P680 that returns part of the excitation energy and is reemitted as light with a peak wavelength at 735 nm. Chlorophyll a fluorescence is also
emitted from PSI and contributes significantly to measurements of minimum
fluorescence (F0), however it does not vary during illumination [85, 86]. Chlorophyll b
in vivo does not emit fluorescence because it transfers the excitation energy with
100% efficiency to chlorophyll a.
Since its discovery, chlorophyll a fluorescence has been extensively used as a
non-invasive way to describe the function of the photosynthetic machinery. Various
methods and equipment have been since developed to make this approach widely
available.
3.3.1 Fast kinetics of fluorescence induction curves, also known as OJIP curves,
are fluorescence transients that are measured upon illumination with a saturating
light pulse of a photosynthetic organism following a period of dark adaptation
(method used in Paper II and V).
Figure 6. Typical chlorophyll a induction transients. Left panel represents an
induction curve of a 15 min dark-adapted Arabidopsis thaliana leaf plotted on a linear
scale. F0 and Fm are here clearly distinguishable. Right panel shows the same
measurement plotted on a logarithmic time scale. Apart from F0 and Fm, sometimes
termed O (origin) and P (peak), the additional J and I intermediate peaks are
observable.
19
Andrei Herdean
Possibly the most widely used parameter derived from induction curves is
Fv/Fm, which is calculated as (Fm-F0)/Fm, and it is generally thought that it indicates
PSII efficiency or PSII quantum yield. However, recent experimental evidence
suggests that fast conformational changes within PSII take place upon illumination
and may account for up to 30% of the fluorescence rise, challenging the notion that
Fv/Fm reflects a true PSII quantum yield [87].
From a typical fluorescence induction curve (Fig. 6) additional parameters can
be derived in what is known as the JIP Test, providing detailed structural, and
functional information about the photosynthetic apparatus, especially PSII [86, 88].
3.3.2 Pulse Amplitude Modulated fluorescence (PAM) is another method that has
been later developed [89], and unlike Fv/Fm it derives its information over longer time
scales (minutes to hours) by determining the Fm’ values, i.e. light-adapted maximum
fluorescence (method used in Papers I to V).
Figure 7. Typical recording of PAM chlorophyll a fluorescence. Plotted data
show a continuous recording of the fluorescence signal of an Arabidopsis thaliana
leaf that has been dark adapted for 30 min prior to the measurement. The
measurement starts with determination of F0 and Fm by exposing the leaf to a brief
800-ms saturating pulse (SP) of 5,000 mol photons m-2 s-1 actinic red light. After the
initial SP the leaf is left without illumination for 1 min to allow the fluorescence signal
to reach F0 levels again, after which the leaf is continuously illuminated with 600
mol photons m-2 s-1 actinic red light. During illumination and the subsequent dark
20
Ion transport in chloroplasts with role in regulation of photosynthesis
adaptation starting at min 11, the leaf is exposed to pre-set saturating pulses to
determine Fm’ values. Red dots next to the time axis indicate when the saturating
pulses have been applied.
From a typical PAM fluorescence measurement (Fig. 7), several useful
parameters can be derived [90]:
 PSII quantum yield (II))
(II) = (Fm’/F)/Fm’
F=

Steady-state fluorescence value prior to the saturation pulse
Electron Transfer Rate at PSII (ETR II) [91, 92]:
ETR (II) =  (II) x PAR x 0.84 x 0.5
PAR = Photosynthetic Active Radiation is the light intensity that the leaf
is exposed to during measurement
0.84 = Percentage of absorbed light (84%) [92]
0.5 = Fraction of the absorbed light distributed to PSI and PSII,
respectively. This value assumes that 50% light is absorbed by
PSII and 50% by PSI

Non-Photochemical Quenching (NPQ) [90]:
NPQ = (Fm-Fm’)/Fm’

Additional parameters such as quantum yield of regulated energy
dissipation (Y(NPQ)), quantum yield of nonregulated energy dissipation
(Y(NO)), coefficient of nonphotochemical quenching (qN), coefficient of
photochemical quenching ((qP)-based on the “puddle” antenna model,
and (qL)- based on the “lake” antenna model) can be calculated using
the same data [93].
 (II) is a parameter that is used to monitor changes in PSII photochemistry in
different experimental setups over longer time intervals. This is a useful indicator
when studying mutants that directly or indirectly affect PSII.
ETR (II) represents a measure of the linear electron flow through PSII and is
determined as mol e- m-2 s-1. Accurate determination of ETR values may become
problematic when comparing wild type (WT) to mutant plants that differ in pigment or
photosynthetic proteins composition because it will alter the two assumed
parameters for light absorption (0.84) and distribution (0.5). These parameters may
vary between young and old plants, and between plant species. For best estimation
of ETR, it is recommended to determine the leaf absorption and PSII/PSI distribution.
21
Andrei Herdean
NPQ monitors the apparent heat loss from PSII, in other words dissipation of
excess energy as heat. Plants can absorb and utilize a certain amount of energy
before light reaches values where it becomes harmful. In order to protect themselves
from photo-damage, chloroplasts dissipate the excess energy as heat [94]. Several
mechanisms work together to dissipate excess energy as a photoprotective
mechanism and are included in NPQ.
NPQ = qE + qZ + qM + qT + qI
qE is the fast energy dependent component, relies on pH, zeaxanthin and
PsbS, and is formed within 10-200 s [95, 96].
qZ, also called zeaxanthin-dependent quencher, is a slow inducible
component that uses pH for de-epoxidation of violaxanthin into zeaxanthin, which is
the pigment responsible for the quenching mechanism. It takes approximately 10-30
min for this mechanism to reach maximum values [95].
qM is a chloroplast photorelocation or avoidance mechanism where
chloroplasts physically rearrange themselves within the cell to shade each other from
excess light [97].
qT is the state transition component that is important under certain light
conditions, specifically under low light, and is dependent on Stn7 –mediated
phosphorylation of LHCII [5]. State transition can be studied as a separate
mechanism in itself using specially designed protocols that monitor steady state
chlorophyll a fluorescence under low light and changes in Fm’ amplitude under the
same conditions [98].
qI is the photoinhibition component, and is ascribed to the photoinhibitory
damage of PSII reaction centers. This component forms slowly, persists for several
hours after illumination has stopped, and is highly dependent on the PSII repair
mechanism [99].
3.4 ElectroChromic band Shift (ECS) at 515 nm was first discovered by Duysens in
1954 [100], and was later demonstrated that it provides a linear measure of the
electric potential across the thylakoid membrane [101]. Because the primary electron
donor and the electron acceptors of the photosynthetic machinery are located on
opposite sides of the thylakoid membrane, as well as the H+ translocating activity of
the Q cycle, an electrochemical H+ gradient is generated across the thylakoid
membrane, also termed as proton motive force (PMF). The electrochemical gradient
has two components: the pH gradient (pH) and the electrical potential gradient
(). Photosynthetic pigments, mainly carotenoids and chlorophyll b, in the presence
of the generated electric field, change their absorption levels and give rise to ECS at
515 nm [102, 103].
There are two general approaches of collecting data using the ECS method,
one using short flashes and recording the 515 signal on a ms time scale (Figure 8),
and another one recording the signal during longer illumination intervals (seconds to
22
Ion transport in chloroplasts with role in regulation of photosynthesis
minutes), as well as transitions from light to dark (Figure 9) (method used in Paper I
to V).
Figure 8. Single turnover flash-induced ECS. 5 min (left panel) or 30 min (middle
panel) dark-adapted Arabidopsis thaliana leaf exposed to a 5-s flash of red actinic
light of 200,000 mol photons m-2 s-1. Right panel shows the information that the
ECS signal contains about the electron transfer chain and ATP synthase.
The types of information that can be obtained using short flashes are the
following:
 PSII and PSI simultaneous charge separation; this information is
gained from the amplitude of the ECS signal upon exposure to a
saturating flash (Figure 8, right panel, marked with green colour). Using
DCMU, the contribution of each separate reaction center can be
established.
 Electron transfer through Cyt b6f; this information is found in the second
slow rise seen in the 30 min dark-adapted leaf in the time range 0 to
~20 ms (Figure 8, right panel, marked with blue colour). In short darkadapted samples, this phase is no longer visible (Figure 8, left panel).
 Membrane conductivity or H+ leakage mainly through ATP synthase;
this information corresponds to the decay of the ECS signal starting
from ~20 ms in the long dark-adapted sample (Figure 8, right panel,
marked with orange) or immediately after the ECS rise after a short
dark-adaptation (Figure 8, left panel). The fast decay of the signal in the
measurement of the 5-min dark-adapted leaf indicates that ATP
synthase activates quickly and can dissipate the H+ gradient, whereas
in the 30-min dark-adapted leaf this process is significantly slower.
Longer recordings of the ECS signal provide distinct type of information from
the short ECS recordings. A dark interval relaxation kinetics (DIRK) analysis of the
ECS signal [104] can give information about the following aspects:
 Relative values for the total PMF. After illumination is stopped, the
ECS signal quickly decays until it reaches a minimum value. The
difference between the steady state during illumination and the new
23
Andrei Herdean



minimum value reached immediately after actinic light is switched off
corresponds to the total PMF, marked with red arrow (Figure 9). This
is also termed as ECST.
ATP synthase H+ conductivity (gH+). When illumination is stopped, the
OEC no longer generates H+, and the PQ pool stops its H+ transport
from chloroplast stroma to thylakoid lumen. Concomitantly, ATP
synthase will continue to function until it completely consumes its
driving force (PMF). This process takes < 1 s and can be calculated
as gH+=1/, where  is the half time of the signal decay (Figure 9
inset).
PMF partitioning into pH (ECSINV) and  (ECSSS) (Figure 9 orange
and blue arrows).
Relative value of steady-state H+ flux (H+). H+ flux is calculated as H+
= PMF × gH+ and linearly correlates with the photosynthetic linear
electron flow [105]. H+ can be considered a quantitative measure of
ATP synthase activity, whereas gH+ a qualitative one.
Due to changes in pigment composition and light scattering during longer
illumination intervals, recordings of the 515 nm signal requires a deconvolution
protocol. When using the WALZ DUAL PAM-100, deconvolution is performed
automatically by simultaneous recording at 550 nm wavelength, and the signal is
directly displayed de-convoluted as P515 (P515 I/Ix103 = 550-515 nm) [102].
Additional deconvolution can be done by simultaneously recording light scattering at
535 nm, which has been recently shown to contribute more than expected to the 515
nm signal [106]. The final wavelength signal will be plotted as 550-515-535 nm.
Furthermore, an additional normalization protocol is used for the PMF value, in order
to compensate for small differences that may occur between biological replicates.
For this purpose, a short 5-s flash-induced ECS (ECSST) recording is made for each
leaf prior to every measurement, and the amplitude of the ECSST is used to
normalize the PMF values between replicates. Corrected PMF = measured PMF ×
(maxECSST / ECSST), where maxECSST is the highest ECSST from all the replicates,
and ECSST is the ECSST corresponding to the PMF that is being normalized.
24
Ion transport in chloroplasts with role in regulation of photosynthesis
Figure 9. Recording of ECS dark interval relaxation kinetics. Recording of a 30min dark-adapted leaf of Arabidopsis thaliana illuminated with 535 mol photons m-2
s-1 actinic red light for 2 min followed by 2 min of darkness. Inset shows the kinetics
of the ECS decay at ms time scale when light is turned off; red line indicates a single
exponential decay fitting of the curve, and the gH+ equation calculated using the
decay half time (). After illumination is stopped, the ECS signal further relaxes and
reaches a new dark baseline within tens of seconds, value that is used to determine
the PMF partitioning as shown with blue () and orange (pH) arrows. Red arrow
indicates PMF size.
25
ROLE OF IONS AND ION TRANSPORTERS
IN PHOTOSYNTHESIS
4
Ion transport in chloroplasts with role in regulation of photosynthesis
4.1 Ion transport mechanisms and regulation of the proton motive force. Ions
play a central role in modulation of PMF size, and partitioning into  and pH.
Counterion movement across the thylakoid membrane functions in order to dissipate
[107-109]. Upon exposure to light, several ion fluxes take place across the
thylakoid membrane: (i) H+ are pumped or generated into the lumen (via PQ and the
OEC) and hence a significant  is generated, (ii) cations (Mg2+, K+, etc.) are
exported from the lumen while anions are imported in order to quickly dissipate part
of the . By reducing  it is expected that PMF partitioning is modulated in such
a way to allow a fast formation of a significant pH [110] There is an increasing
number of mutants defective in thylakoid ion fluxes which are unable to reduce
and indeed cannot form a significant pH ([33], Paper IV and V). Size of total
PMF, known to regulate ATP synthase activity [111], can also be modulated by
changes in ion homeostasis (Paper II to V). Below, we discuss the specific role of
each of the major ions in the chloroplast in regulation of PMF across the thylakoid
membrane.
4.1.1 Potassium (K+) is the second most abundant ion in the chloroplast, second
only to H+. Recent publications [32, 33] and Paper I have demonstrated that K+ plays
a crucial role in regulation of the PMF by altering the pH /  stoichiometry. Two
types of transport proteins have been found in the thylakoid membrane of
Arabidopsis that mediate K+ fluxes (Figure 10).
TPK3 is a two-pore K+ channel, shown to regulate photosynthesis by
exporting K+ from the lumen to allow translocation of more H+ into the lumen and
formation of a significant pH. Knockout mutants displayed a two fold decrease in
pH, and consequently a large increase in  without any changes in PMF size [33].
The physiological effects of this alteration in trans-thylakoid energy storage, when
studied in knockout mutants, resulted in reduced plant growth and hypersensitivity to
high light due to impaired photoprotective mechanisms (NPQ) dependent on pH.
KEA3 is the second characterized protein ([32], Paper I) that regulates K+
fluxes across the thylakoid membrane. It functions as a K+/H+ antiporter, and unlike
TPK3, it is proposed to load K+ into the lumen by using the pH formed during
illumination. KEA3 has a KTN domain that may be regulated by the redox state of the
chloroplast stroma, and hence may function as part of a PSI feedback mechanisms
[32]. Arabidopsis thaliana knockout mutants of KEA3 do not have a growth
phenotype but the pH /  stoichiometry is altered in favour of pH, with no
changes in PMF size. Its major role has been shown to be under changing light
conditions, when transitions from high light to low light occur, as well as transitions
from dark to low light.
Together TPK3 and KEA3 form an elegant mechanism for regulating K+ fluxes
across the thylakoid membrane, and indirectly adjusting photosynthesis to the light
environment by altering the PMF composition.
29
Andrei Herdean
Figure 10. Dual protein mechanism for K+ cycling in the chloroplast. Schematic
representation of TPK3 and KEA3 regulated K+ fluxes across the thylakoid
membrane (left image). A similar mechanism functions across the chloroplast inner
envelope where KEA1& KEA2 export K+ to the cytosol (right image). A chloroplast
K+-loading mechanism is still missing.
KEA1 and KEA2, additional members of the KEA family, have been localized
to the chloroplast inner envelope (Paper I) and function in a H+-driven K+ export
system from chloroplast stroma to the cytosol. While these proteins directly regulate
the electrochemical gradient across the envelope membrane they do have a
downstream effect on the thylakoid membrane. Knockout of either KEA1 or KEA2 did
not have a significant impact on plant growth or photosynthesis, however, the
knockout of both proteins resulted in a severely stunted growth, disrupted
chloroplasts, altered PMF size and partitioning (Paper I).
4.1.2 Calcium (Ca2+) is found in the chloroplast at ten-fold lower concentrations
than K+ [112], however, it may still play an important role in size and composition of
PMF. In Paper II we show that the Arabidopsis photosynthesis-affected mutant71
(pam71) has an altered Ca2+ homeostasis in the chloroplast. The pam71 mutant
accumulates Ca2+ inside the thylakoid lumen, which results in a two fold increase in
PMF size, altered  / pH stoichiometry due to significantly increased  as well
as reduced ATP synthase conductivity. TPK3 is Ca2+ regulated [33], and our data
cannot completely exclude the possibility that the observed phenotype in PMF is due
to inhibition of TPK3. The altered Ca2+ distribution within the chloroplast of pam71
may be caused by a defective Ca2+ efflux activity in the thylakoid membrane, for
which the protein responsible remains to be identified.
4.1.3 Phosphate (Pi) has been proposed to be present in the thylakoid lumen as a
component of the lumen-located nucleotide metabolism [113, 114], hence its
presence in the thylakoid lumen may contribute to PMF modulation. The current
model for import and export of phosphate does not suggest that it is a fast acting
mechanism evolved specifically for charge compensation, but rather for the
nucleotide-dependent mechanisms. Two proteins are known to be localized to the
thylakoid membrane that are part of the phosphate transport mechanism: the
thylakoid ATP/ADP carrier (TAAC) [43, 115] and the thylakoid Pi transporter PHT4;1
30
Ion transport in chloroplasts with role in regulation of photosynthesis
[46, 47] (Fig. 11). PHT4;1 has been found by other studies in the chloroplast inner
envelope [48, 49]. If this is the case, then a lumenal phosphate export mechanism
still remains elusive (Fig. 11).
In Paper III, we show that PHT4;1 knockout mutants have a transient increase
in pH, and consequently a significant decrease in , whereas the total PMF is not
affected. Furthermore, thylakoid membrane conductivity (gH+) decreases up to 50%
in these mutants as compared to WT, a phenomenon that was previously reported
under conditions of Pi depletion in the stroma [105]. The observation in Paper III that
addition of Pi during growth of the PHT4;1 mutants restores the WT gH+ supports the
idea of Pi limitation for ATP synthase in the stroma of the mutants. In addition, the
reduced growth phenotype of the mutants was suppressed by high phosphate.
Another previously reported effect of stromal Pi depletion is a significant increase in
PMF size without changing the partitioning [105] which is not observed in PHT4;1
mutants that instead have a transiently increased pH contribution to PMF (Paper
III). This difference can be explained by an accumulation of Pi in the lumen of the
mutants (due to defective Pi export), that dissipates part of , and hence keeps
PMF size at levels similar to wild-type. The transient nature of this effect is probably
due to the fact that the lumenal Pi concentration is significant for membrane
depolarization only before the import of other major anions is initiated (e.g., Cl-), after
which lumenal Pi concentration becomes small relative to that of other anions. Taken
together, the phenotype of PHT4;1 knockout mutants support the location of PHT4;1
in the thylakoid membrane, where it acts as a local supplier of Pi from the thylakoid
lumen to the ATP synthase.
In contrast to PHT4;1, knockout mutants of envelope located PHT2;1 [51, 52,
116], PHT4;4 [53, 54], PHT4;5 [53] or TPT [53, 117] do not have an apparent growth
phenotype or altered photosynthetic performance. It is conceivable that envelopelocalized Pi transporters may also alter PMF size and partitioning through a
downstream effect by limiting the Pi supply to ATP synthase, however no
experimental data is available to confirm this. Interestingly, PHT4;4 has been shown
using in vitro assays to be able to transport ascorbate, in addition to Pi into the
chloroplast [54]. Authors of the same paper predicted a similar function for PHT4;1 in
the thylakoid membrane. However, this possibility is excluded as based on our
finding that the PHT4;1 knockout mutants do not display altered ascorbate content in
the thylakoid lumen (Paper III).
31
Andrei Herdean
Figure 11. Phosphate cycling across the thylakoid membrane. Blue arrows
indicate Pi pathway in the form of nucleoside triphosphates (ATP and GTP) and in
free form after GTP hydrolysis by the PsbO extrinsic subunit of PSII. Following import
into the lumen via TAAC, ATP is converted to GTP by nucleoside diphosphate kinase
3 (NDPK3). GTP is bound and hydrolysed by PsbO in a process associated with D1
turnover [114, 118], releasing free Pi into the lumen. In the thylakoid-location
scenario (left image), PHT4;1 transports Pi back to the chloroplast stroma, where it is
used as substrate by the ATP synthase. If however PHT4;1 is located to the
chloroplast inner envelope (right image), it will simply act as a Pi chloroplast import
system together with other envelope transporters (PHT2;1, PHT4;4 and triose-Pi/Pi
transporter). This second possibility however still requires a protein to export Pi from
the lumen to the stroma.
32
Ion transport in chloroplasts with role in regulation of photosynthesis
4.1.4 Chloride (Cl−) is the most abundant anion present in the chloroplast and is
thought to be important for membrane depolarization. A thylakoid-located Cl− channel
activity has been first reported using a patch-clamp approach on isolated thylakoids
from Peperomia metallica [109]. A member of the CLC chloride family, CLCe, has
been localized to the thylakoid membrane [35, 36], and has been long considered to
directly regulate Cl− or NO3− ion fluxes across thylakoids. In Paper IV, we identify a
second thylakoid-located Cl− channel in Arabidopsis, VCCN1. ECS measurements of
CLCe knockout mutants (Paper V) show small but statistically significant changes in
PMF size,  / pH stoichiometry, as well as ATP synthase activity. A similar but
greater PMF phenotype was observed in knockout mutants of VCCN1 (Paper IV).
The results from ECS measurements confirm that both CLCe and VCCN1 play role
in depolarization of the thylakoid membrane.
Despite the identification of two distinct Cl−-translocating proteins in
thylakoids, VCCN1 and CLCe (the transport function of the latter remains to be
confirmed by heterologous expression), the complete mechanism for Cl- fluxes has
not yet been solved. It is however established that a Cl− influx to the lumen takes
place in the light, and an efflux in darkness [119]. Four hypothetical models are
presented in Figure 12 for how these fluxes may take place in the context of VCCN1
and CLCe. Several other possibilities may exist and could also be considered. In
order to elucidate this mechanism, more experimental data are required about the
type of transport process that CLCe performs (channel versus transporter) as well as
the substrate specificity, and a more accurate determination of the lumenal and
stromal Cl− concentrations in the light and in the dark.
Figure 12. Hypothetical models of Cl− transport across the thylakoid
membrane. (I) VCCN1 and CLCe are driven by  and both transport Cl− in and out
of the lumen, but are differently regulated or have different levels of specificity.
Export can take place immediately after light is switched off due to change in
electrical polarity across the membrane. (II) VCCN1 is driven in the same way by 
for import and export, whereas CLCe uses pH to drive import of Cl− against its
concentration gradient. (III) VCCN1 imports Cl- in light but does not perform export
(the inverted polarity in transition to darkness may not be energy sufficient), whereas
CLCe uses pH upon transition to darkness for Cl− efflux. (IV) Both proteins use 
33
Andrei Herdean
but are differently gated and VCCN1 provides the influx, whereas CLCe the efflux of
Cl−.
4.1.5 Other ions like Mg2+, which is present in mM concentrations in the
chloroplast, Mn2+, Zn2+, Cu2+, Fe2+, NO3− or NO2− have been less studied in the
context of their role in regulation of the PMF. Nevertheless, they may contribute
directly or indirectly to the overall electrochemical gradient across the thylakoid
membrane. The proteins responsible for mediating these ion fluxes remain largely
elusive.
4.2 Regulation of electron transfer.
Arguably the most important component of the ETC, that is dependent on a
specific configuration of ions, is the OEC. The assembly of the Mn4CaO5 inorganic
cluster at the donor side of PSII is not fully understood, but it is obvious that during
the assembly process Ca2+ and Mn2+ are strictly required in the thylakoid lumen. Clions have also been found in the vicinity of the Mn4CaO5 cluster, and may have a
role in H+ channelling from the core of the OEC after oxidation of water [11, 120,
121].
The Arabidopsis pam71 mutant has a significant loss of PSII activity due to
impaired and unstable OEC, and resembles a double psbo1×psbo2 mutant (Paper
II). Since pam71 accumulates Ca2+ inside the thylakoid lumen, it is plausible that this
causes the observed defect at OEC site. This possibility is supported by the notion
that PsbO, known to stabilize the OEC, is washed off the PSII complex in the
presence of excess Ca2+ [122]. This finding emphasizes the importance of Ca2+
homeostasis for the optimal function of OEC and entire ETC.
Although chloride is known to have a role in oxygen evolution, knockout
mutants of VCCN1 or CLCe do not seem to have a significant impact on the OEC
(Paper IV and V).
PSII, Cyt b6f and PSI have iron clusters that directly participate in transfer of
−
e . Approximately 80% of cellular Fe2+ is stored in the chloroplast [38], and under
Fe2+ starvation plants undergo chlorosis. Fe2+ transport in to the chloroplast takes
places via several transporter proteins: MAR1 [57], NAP14 [58], YLS4 [59, 60], YLS6
[59, 60], PIC1 [61, 62], and NiCo [61]. No Fe2+ transport mechanism or requirement
has been proposed across the thylakoid membrane. Cu2+ is a redox cofactor for PC
[123], hence is directly required for electron transfer between Cyt b6f and PSI. Cu2+
import to the chloroplast is proposed to take place via HMA6 [66, 68] and HMA1,
whereas import to the thylakoid lumen via HMA8 [48, 66].
4.3 Regulation of thylakoid architecture.
Regulation of thylakoid stacking and unstacking is an important process for
PSII repair [3], state transition [124], NPQ [125] and spill-over of excitation energy
[126]. Two distinct but interconnected phenomena govern the thylakoid stacking and
34
Ion transport in chloroplasts with role in regulation of photosynthesis
unstacking: electrostatic screening of the negative charges from membrane bound
proteins at the surface of the membrane [127, 128], and increased osmotic pressure
in the lumen [3]. -driven influx of Cl− in the lumen increases the osmotic potential,
hence induces lumen swelling [119]. Knockout mutants of the thylakoid chloride
channel VCCN1 have altered thylakoid structures, confirming the role of Clhomeostasis in regulation of thylakoid ultrastructure (Paper IV). Mg2+ is well known
to be a catalyst for grana stacking [129, 130], whereas Ca2+, Mn2+ and K+ are known
to be less effective in this process [131]. Nevertheless, knockout mutants of the
thylakoid potassium channel TPK3 have an altered ultrastructure under certain light
intensities [33].
4.4 Signalling and enzymatic activation.
Calcium is predominantly bound to the negatively charged thylakoid
membrane or to calcium-binding proteins, and only a small fraction is free resting.
Light-dependent Ca2+ influx into the lumen has been associated with Mg2+ efflux to
the stroma [132, 133]. Ca2+ is involved in a wide range of regulatory processes in the
chloroplast, by either direct binding1 of Ca2+, Ca2+ dependent phosphorylation2, or
calmodulin (CaM) binding3:
 linear electron flow (PsbO1)
 cyclic electron flow (NDH-F2)
 state transition (PsaH2)
 PSII repair (FtsH22 and 52)
 carbon fixation (FBPase, SBPase, TKL2)
 transporters and channels gating (TPK32, TIC1102, TIC323)
 regulation of stomata function and photoaclimation (CAM1-3) [112].
Copper is required in the thylakoid lumen for PC maturation [123], and
additionally it regulates its thylakoid loading mechanism by binding to HMA8 (ATPdependent Cu2+ transporter) to initiate its degradation by Clp proteases [134]. Cu2+,
Zn2+ and Fe2+ are required for the activity of superoxide dismutase, a scavenger of
reactive oxygen species [135].
Zinc has been shown to stimulate the proteolytic activity of FtsH2, and to
promote specific degradation of Rieske FeS protein of the Cyt b6f [136, 137].
Magnesium catalyses enzymatic reactions of acetohydroxy acid
isomeroreductase that is involved in biosynthesis of the amino acids isoleucine,
valine, and leucine [138]. The ATP synthase CF1 subunit also binds Mg2+ and
becomes reversibly inactivated [139].
35
Andrei Herdean
4.5 Regulation of photoprotective mechanisms.
In natural environments plants absorb more energy from light that can be used
for photochemistry, and in order to avoid photodamage the excess energy is
dissipated as heat in a process called non-photochemical quenching (NPQ). NPQ
incorporates several mechanisms that are directly or indirectly dependent on pH
(See section 3.3.2).
Knockout mutants of chloroplast thylakoid and inner envelope ion transporters
have almost always an altered NPQ [32, 33, 54] (Paper I, III, IV, and V). However,
this effect cannot be attributed to the transporters themselves, since they do not
directly participate in NPQ, but to a secondary effect of the altered pH (in the case
of TPK3, KEA1-3, PHT4;1 and VCCN1 in Papers I, III and IV), impaired enzymatic
activity of the xanthophyll cycle (PHT4;4, [54]) or possible changes in thylakoid
architecture (CLCe in Paper V). PHT4;1 changes the kinetics of NPQ by altering Pi
availability to ATP synthase, which most likely results in a faster acidification of the
thylakoid lumen that in turn affects NPQ (Paper III).
In nature, plants experience rapid fluctuations in light intensity that change up
to 100 fold within seconds to minutes [140]. In order to avoid photodamage, plants
can modulate the response time of NPQ by quickly adjusting the pH. Previously
KEA3 has been shown to modulate NPQ upon rapid shifts from high to low light [32].
In Paper IV we show that VCCN1 is able to modulate the NPQ response time upon
rapid shifts from low to high light.
36
CONCLUSIONS AND OUTLOOK
5
Ion transport in chloroplasts with role in regulation of photosynthesis
The work presented in this thesis answers several questions about the role in
regulation of photosynthesis of a number of ion channels and transporters from the
thylakoid membrane and chloroplast inner envelope of Arabidopsis.
Regulation of K+ transport in chloroplasts is presented as a complete
mechanism governed by KEA3 and TPK3 across the thylakoid membrane and
partially by KEA1 and KEA2 across the inner envelope (Paper I). The cytosol to
stroma import mechanism still remains to be resolved. Nevertheless, the importance
of KEA1, KEA2 and KEA3 for plant growth is studied, and their role in K+
homeostasis, regulation of PMF and photoprotection is presented in Paper I.
Ca2+ transport across the thylakoid membrane has been demonstrated since
1999 [133], however the protein(s) responsible for these fluxes have not been
identified. In Paper II, a mutant affected in Ca2+ homeostasis is identified (pam71).
This alteration is shown to have implications for PSII function and regulation of PMF.
A defective activity for Ca2+ efflux from the thylakoid lumen may be responsible for
the observed phenotype, for which the responsible protein remains to be identified.
In Paper III, role of the PHT4;1 in phosphate transport across the thylakoid
membrane is addressed. We present a new model for Pi metabolism where PHT4;1
is a localized supplier of Pi to ATP synthase. This discovery has implications for plant
growth,  / pH stoichiometry and photoprotection.
In Paper IV a novel voltage-dependent Cl− channel (VCCN1) is identified in
thylakoids. VCCN1 is the first confirmed (by electrophysiology) Cl- channel from the
thylakoid membrane. Cl− fluxes in the chloroplast have long been thought to regulate
membrane depolarization, and to induce changes in thylakoid ultrastructure. These
predictions are confirmed in Paper IV, and additionally we show that PMF-driven Cl−
fluxes also regulate formation of NPQ upon rapid shifts from low to high light.
Before the discovery of VCCN1 in the thylakoids, another protein from the
CLC family (CLCe) has been localized to the same membrane, and was thought to
mediate Cl− fluxes. The Cl− transport function of CLCe has not yet been confirmed by
heterologous expression. However, in Paper V its role in photosynthesis is
investigated, and similarly to VCCN1, it was found to regulate membrane
depolarization but to a smaller extent. Additionally, a specific function in state
transition and regulation of the electron transfer was found.
Taken together, the findings presented in this thesis contribute to the
increasing picture of ion channels and transporters from the chloroplast and their
importance in photosynthesis. Furthermore, studies of the possible interactions
between these proteins will help to better describe the network of ion fluxes in the
chloroplast.
39
Andrei Herdean
Due to their role in modulation of PMF and its partitioning, ion transporters
could potentially be used for engineering photosynthetic organisms with maximized
light use efficiency, and shortened response time of photoprotective mechanisms.
VCCN1 together with KEA3 may be used to accelerate or slow down the response
time of NPQ upon fast changes in light intensity. CLCe on the other hand may be
used to modulate the response time for changes in light quality. Several other
features of transporters and channels may be further employed for optimization or
improvement of photosynthesis.
40
ACKNOWLEDGEMENTS
6
I would like to thank Cornelia for guiding me through my PhD, being my scientific
mentor, and for all the support throughout these four years. I greatly appreciate supporting
me to test new ideas and experiments, and for teaching me how to do proper science. I
would not have made it to where I am today without your help.
Thank you, Björn, for being more than my co-supervisor, and for making the lab a
pleasant and fun environment. Your enthusiasm for scientific ”uncharted territory” I
found it unparalleled.
I also thank my examiner, Adrian, for making sure I was on the right track during
my PhD studies, and for all the helpful advices.
Many thanks to my current and former lab colleagues: Lisa, Hugues, Somnath,
Milton, Lan, Azeez, and Jenny. I have learned many things from our talks about science
and not only.
I thank the co-authors of the papers I am included in for giving me the opportunity
to work and learn with you: Henning H.K., Anja S., Milton K., Lisa A., Azeez B., Hugues
N., Győző G., Ottó Z., Gergely N., Renáta Ü., Ildikó S., Henrik A., Wayne V. K., Anders N.,
Oskar J., Katalin S, Lan Y., Rikard F., Pádraic F., Benoît S, Mats XA..
Thank you, Ingela D., Henrik A., Sven T., and Ylva H., for making the department
a fun, pleasant and dynamic place to be part of.
I also thank all the PhD students from our department. I had a lot of fun being part
of this cohort. May all your papers be accepted!
De asemenea le mulțumesc părinților mei pentru fundația solidă cu care m-au
trimis în lume, și pentru toți anii de acasă care m-au facut cine sunt eu azi.
Egy bölcs ember egyszer azt mondta: "minden erős férfi mögött áll egy még erősebb
nő". Hiszem, hogy ez így igaz, és köszönöm neked Ági, mert te vagy életem
legcsodálatosabb része!
43
REFERENCES
7
Ion transport in chloroplasts with role in regulation of photosynthesis
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
Olson, J.M., et al., Thinking about the evolution of photosynthesis. Photosynthesis Research,
2004. 80(1‐3): p. 373‐386.
Holland, H.D., The oxygenation of the atmosphere and oceans. Philosophical Transactions of
the Royal Society B: Biological Sciences, 2006. 361(1470): p. 903‐915.
Kirchhoff, H., et al., Dynamic control of protein diffusion within the granal thylakoid lumen.
Proceedings of the National Academy of Sciences, 2011. 108(50): p. 20248‐20253.
Kowalczyk, N., et al., Photosynthesis in Chondrus crispus: The contribution of energy spill‐over
in the regulation of excitonic flux. Biochimica et Biophysica Acta (BBA) ‐ Bioenergetics, 2013.
1827(7): p. 834‐842.
Mikko, T., et al., State transitions revisited—a buffering system for dynamic low light
acclimation of Arabidopsis. Plant molecular biology, 2006. 62(4‐5): p. 779‐793.
Vinyard, D.J., et al., Photosystem II: the reaction center of oxygenic photosynthesis. Annual
review of biochemistry, 2013. 82: p. 577‐606.
Joliot, P., et al., Un nouveau modele des centres photochimiques du systeme ii.
Photochemistry and Photobiology, 1969. 10(5): p. 309‐329.
Kok, B., et al., Cooperation of charges in photosynthetic O2 evolution–I. A linear four step
mechanism. Photochemistry and Photobiology, 1970. 11(6): p. 457‐475.
Forbush, B., et al., Cooperation of charges in photosynthetic O2 evolution‐II. damping of
flash yield oscillation, deactivation. Photochemistry and Photobiology, 1971. 14(3): p. 307‐
321. Cox, N., et al., Electronic structure of the oxygen‐evolving complex in photosystem II prior to
OO bond formation. Science, 2014. 345(6198): p. 804‐808.
Umena, Y., et al., Crystal structure of oxygen‐evolving photosystem II at a resolution of 1.9 Å.
Nature, 2011. 473(7345): p. 55‐60.
Kern, J., et al., Simultaneous femtosecond X‐ray spectroscopy and diffraction of photosystem
II at room temperature. Science, 2013. 340(6131): p. 491‐495.
Whitelegge, J.P., et al., Full subunit coverage liquid chromatography electrospray ionization
mass spectrometry (LCMS+) of an oligomeric membrane protein ‐ Cytochrome b6f complex
from spinach and the cyanobacterium Mastigocladus laminosus. Molecular & Cellular
Proteomics, 2002. 1(10): p. 816‐827.
Cramer, W.A., et al., Transmembrane traffic in the cytochrome b6f complex. Annu. Rev.
Biochem., 2006. 75: p. 769‐790.
Munekage, Y., et al., Cyclic electron flow around photosystem I is essential for
photosynthesis. Nature, 2004. 429(6991): p. 579‐582.
Zhang, H., et al., Ferredoxin:NADP+ oxidoreductase is a subunit of the chloroplast cytochrome
b6 f complex. Journal of Biological Chemistry, 2001. 276(41): p. 38159‐38165.
Fromme, P., et al., Unraveling the photosystem I reaction center: a history, or the sum of
many efforts. Photosynthesis Research, 2004. 80(1‐3): p. 109‐124.
Rögner, M., et al., Mono‐, di‐and trimeric PS I reaction center complexes isolated from the
thermophilic cyanobacterium Synechococcus sp.: Size, shape and activity. Biochimica et
Biophysica Acta (BBA)‐Bioenergetics, 1990. 1015(3): p. 415‐424.
Boekema, E., et al., Evidence for a trimeric organization of the photosystem I complex from
the thermophilic cyanobacterium Synechococcus sp. FEBS letters, 1987. 217(2): p. 283‐286.
Minagawa, J., State transitions—The molecular remodeling of photosynthetic
supercomplexes that controls energy flow in the chloroplast. Biochimica et Biophysica Acta
(BBA) ‐ Bioenergetics, 2011. 1807(8): p. 897‐905.
Süss, K.‐H., et al., Evidence for an α 3, β 3, γ, δ, I, II, ε, III 5 subunit stoichiometry of
chloroplast ATP synthetase complex (CF 1− CF 0). FEBS letters, 1982. 144(2): p. 213‐218.
Poetsch, A., et al., Biophysics and bioinformatics reveal structural differences of the two
peripheral stalk subunits in chloroplast ATP synthase. Journal of biochemistry, 2007. 141(3):
p. 411‐420.
47
Andrei Herdean
23.
24.
25.
26.
27.
28.
29.
30.
31.
32.
33.
34.
35.
36.
37.
38.
39.
40.
41.
42.
43.
44.
45.
48
Fromme, P., et al., Isolation and characterization of a supramolecular complex of subunit III
of the ATP‐synthase from chloroplasts. Zeitschrift für Naturforschung C, 1987. 42(11‐12): p.
1239‐1245.
Dodd, A., et al., The circadian regulation of photosynthesis. Photosynthesis Research, 2014.
119(1‐2): p. 181‐190.
Long, S.P., et al., Photoinhibition of photosynthesis in nature. Annual Review of Plant
Physiology and Plant Molecular Biology, 1994. 45(1): p. 633‐662.
Scheller, H., et al., Photoinhibition of photosystem I. Planta, 2005. 221(1): p. 5‐8.
Ruban, A.V., et al., The photoprotective molecular switch in the photosystem II antenna.
Biochimica et Biophysica Acta (BBA) ‐ Bioenergetics, 2012. 1817(1): p. 167‐181.
Müller, P., et al., Non‐photochemical quenching. a response to excess light energy. Plant
physiology, 2001. 125(4): p. 1558‐1566.
Li, X.‐P., et al., Regulation of photosynthetic light harvesting involves intrathylakoid lumen ph
sensing by the PsbS protein. Journal of Biological Chemistry, 2004. 279(22): p. 22866‐22874.
Niyogi, K.K., et al., Arabidopsis mutants define a central role for the xanthophyll cycle in the
regulation of photosynthetic energy conversion. The Plant Cell, 1998. 10(7): p. 1121‐1134.
Johnson, M.P., et al., Photoprotective energy dissipation involves the reorganization of
photosystem II light‐harvesting complexes in the grana membranes of spinach chloroplasts.
The Plant Cell, 2011. 23(4): p. 1468‐1479.
Armbruster, U., et al., Ion antiport accelerates photosynthetic acclimation in fluctuating light
environments. Nature communications, 2014. 5.
Carraretto, L., et al., A thylakoid‐located two‐pore K+ channel controls photosynthetic light
utilization in plants. Science, 2013. 342(6154): p. 114‐118.
Saier, M.H., et al., The Transporter Classification Database. Nucleic Acids Research, 2014.
42(D1): p. D251‐D258.
Marmagne, A., et al., Two members of the Arabidopsis CLC (chloride channel) family, AtCLCe
and AtCLCf, are associated with thylakoid and Golgi membranes, respectively. Journal of
Experimental Botany, 2007: p. 3385‐3393.
Lv, Q.‐d., et al., Cloning and molecular analyses of the Arabidopsis thaliana chloride channel
gene family. Plant Science, 2009. 176(5): p. 650‐661.
Cornelia Spetea, B.S., Photosynthesis and light stress in a model plant: role of chloroplast
transporters, in Handbook of plant and crop stress, M. Pessarakli, Editor. 2010, CRC Press:
USA. p. 361‐389.
Finazzi, G., et al., Ions channels/transporters and chloroplast regulation. Cell calcium, 2014.
Haswell, E.S., et al., MscS‐like proteins control plastid size and shape in Arabidopsis thaliana.
Current Biology, 2006. 16(1): p. 1‐11.
Teardo, E., et al., Dual localization of plant glutamate receptor AtGLR3.4 to plastids and
plasmamembrane. Biochimica et Biophysica Acta (BBA)‐Bioenergetics, 2011. 1807(3): p. 359‐
67.
Drummond, R.S.M., et al., A putative magnesium transporter AtMRS2‐11 is localized to the
plant chloroplast envelope membrane system. Plant Science, 2006. 170(1): p. 78‐89.
Thuswaldner, S., et al., Identification, expression, and functional analyses of a thylakoid
ATP/ADP carrier from arabidopsis. Journal of Biological Chemistry, 2007. 282(12): p. 8848‐
8859. Yin, L., et al., Role of thylakoid ATP/ADP carrier in photoinhibition and photoprotection of
photosystem II in Arabidopsis. Plant physiology, 2010. 153(2): p. 666‐677.
Gigolashvili, T., et al., The Arabidopsis thylakoid ADP/ATP carrier TAAC has an additional role
in supplying plastidic phosphoadenosine 5'‐phosphosulfate to the cytosol. Plant Cell, 2012.
24(10): p. 4187‐204.
Han, L., et al., Identification and localized expression of putative K+/H+ antiporter genes in
Arabidopsis. Acta Physiologiae Plantarum, 2015. 37(5): p. 1‐14.
Ion transport in chloroplasts with role in regulation of photosynthesis
46.
47.
48.
49.
50.
51.
52.
53.
54.
55.
56.
57.
58.
59.
60.
61.
62.
63.
64.
65.
Pavón, L.R., et al., Arabidopsis ANTR1 Is a thylakoid Na+‐dependent phosphate transporter
functional characterization in escherichia coli. Journal of Biological Chemistry, 2008. 283(20):
p. 13520‐13527.
Yin, L., et al., The membrane proteome of stroma thylakoids from Arabidopsis thaliana
studied by successive in‐solution and in‐gel digestion. Physiologia plantarum, 2015.
154(3): p. 433‐446.
Tomizioli, M., et al., Deciphering thylakoid sub‐compartments using a mass spectrometry‐
based approach. Molecular & Cellular Proteomics, 2014. 13(8): p. 2147‐2167.
Ferro, M., et al., AT_CHLORO, a comprehensive chloroplast proteome database with
subplastidial localization and curated information on envelope proteins. Molecular & Cellular
Proteomics, 2010. 9(6): p. 1063‐1084.
Aranda‐Sicilia, M.N., et al., Arabidopsis KEA2, a homolog of bacterial KefC, encodes a K+/H+
antiporter with a chloroplast transit peptide. Biochimica et Biophysica Acta (BBA)‐
Biomembranes, 2012. 1818(9): p. 2362‐2371.
Versaw, W.K., et al., A chloroplast phosphate transporter, PHT2;1, influences allocation of
phosphate within the plant and phosphate‐starvation responses. The Plant Cell, 2002. 14(8):
p. 1751‐1766.
Ferro, M., et al., Integral membrane proteins of the chloroplast envelope: Identification and
subcellular localization of new transporters. Proceedings of the National Academy of
Sciences, 2002. 99(17): p. 11487‐11492.
Guo, B., et al., Functional analysis of the Arabidopsis PHT4 family of intracellular phosphate
transporters. New Phytologist, 2008. 177(4): p. 889‐898.
Miyaji, T., et al., AtPHT4;4 is a chloroplast‐localized ascorbate transporter in Arabidopsis.
Nature communications, 2015. 6.
Sugiura, M., et al., A nitrite transporter associated with nitrite uptake by higher plant
chloroplasts. Plant and cell physiology, 2007. 48(7): p. 1022‐1035.
Cao, M.J., et al., SULTR3;1 is a chloroplast‐localized sulfate transporter in Arabidopsis
thaliana. The Plant Journal, 2013. 73(4): p. 607‐616.
Conte, S., et al., Multiple antibiotic resistance in Arabidopsis is conferred by mutations in a
chloroplast‐localized transport protein. Plant physiology, 2009. 151(2): p. 559‐573.
SHIMONI‐SHOR, E., et al., Disruption of Nap14, a plastid‐localized non‐intrinsic ABC
protein in Arabidopsis thaliana results in the over‐accumulation of transition metals and in
aberrant chloroplast structures. Plant, cell & environment, 2010. 33(6): p. 1029‐1038.
Divol, F., et al., The Arabidopsis Yellow Stripe LIKE4 and 6 transporters control iron release
from the chloroplast. The Plant Cell, 2013. 25(3): p. 1040‐1055.
Conte, S., et al., Arabidopsis thaliana Yellow Stripe1‐Like4 and Yellow Stripe1‐Like6 localize to
internal cellular membranes and are involved in metal ion homeostasis. Frontiers in plant
science, 2013. 4.
Duy, D., et al., PIC1, an ancient permease in Arabidopsis chloroplasts, mediates iron
transport. The Plant Cell, 2007. 19(3): p. 986‐1006.
Duy, D., et al., The chloroplast permease PIC1 regulates plant growth and development by
directing homeostasis and transport of iron. Plant physiology, 2011. 155(4): p. 1709‐1722.
Müller, M., et al., Decreased capacity for sodium export out of Arabidopsis chloroplasts
impairs salt tolerance, photosynthesis and plant performance. The Plant Journal, 2014. 78(4):
p. 646‐658.
Winget, G.D., et al., Formation of ATP by the adenosine triphosphatase complex from spinach
chloroplasts reconstituted together with bacteriorhodopsin. Biochimica et Biophysica Acta
(BBA) ‐ Bioenergetics, 1977. 460(3): p. 490‐499.
Pick, U., et al., Purification and reconstitution of the N,N'‐dicyclohexylcarbodiimide‐sensitive
ATPase complex from spinach chloroplasts. Journal of Biological Chemistry, 1979. 254(8): p.
2793‐2799.
49
Andrei Herdean
66.
67.
68.
69.
70.
71.
72.
73.
74.
75.
76.
77.
78.
79.
80.
81.
82.
83.
84.
85.
86.
87.
88.
50
Abdel‐Ghany, S.E., et al., Two P‐type ATPases are required for copper delivery in Arabidopsis
thaliana chloroplasts. Plant Cell, 2005. 17(4): p. 1233‐51.
Seigneurin‐Berny, D., et al., HMA1, a new Cu‐atpase of the chloro plast envelope, is essential
for growth under adverse light conditions. Journal of Biological Chemistry, 2006. 281(5): p.
2882‐2892.
Shikanai, T., et al., PAA1, a P‐type ATPase of Arabidopsis, functions in copper transport in
chloroplasts. The Plant Cell, 2003. 15(6): p. 1333‐1346.
Huang, L., et al., Characterization of a gene encoding a Ca2+‐ATPase‐like protein in the plastid
envelope. Proceedings of the National Academy of Sciences, 1993. 90(21): p. 10066‐10070.
Laibach, F., Zur Frage nach der individualität der Chromosomen im Pflanzenreich (Doctoral
dissertation). 1907.
Laibach, F., Arabidopsis thaliana (L.) Heynh. als Objekt für genetische und
entwicklungsphysiologische Untersuchungen. Botanical Archives, 1943. 44: p. 439‐455.
Lloyd, A.M., et al., Transformation of Arabidopsis thaliana with Agrobacterium tumefaciens.
Science, 1986. 234(4775): p. 464‐466.
Chang, C., et al., Molecular cloning and DNA sequence of the Arabidopsis thaliana alcohol
dehydrogenase gene. Proceedings of the National Academy of Sciences, 1986. 83(5): p. 1408‐
1412. Arabidopsis Genome Initiative, Analysis of the genome sequence of the flowering plant Arabidopsis thaliana. Nature, 2000. 408(6814): p. 796.
Krysan, P.J., et al., T‐DNA as an insertional mutagen in Arabidopsis. The Plant Cell, 1999.
11(12): p. 2283‐2290.
Bruce, B.D., Chloroplast transit peptides: structure, function and evolution. Trends in Cell
Biology, 2000. 10(10): p. 440‐7.
Schröder, W.P., et al., Update on chloroplast proteomics. Photosynthesis Research, 2003.
78(3): p. 181‐193.
Ferro, M., et al., Proteomics of the chloroplast envelope membranes from Arabidopsis
thaliana. Molecular & Cellular Proteomics, 2003. 2(5): p. 325‐345.
van Wijk, K.J., Plastid proteomics. Plant Physiology and Biochemistry, 2004. 42(12): p. 963‐
977. Friso, G., et al., In‐depth analysis of the thylakoid membrane proteome of Arabidopsis
thaliana chloroplasts: new proteins, new functions, and a plastid proteome database. The
Plant Cell, 2004. 16(2): p. 478‐499.
Kleffmann, T., et al., plprot: A comprehensive proteome database for different plastid types.
Plant and cell physiology, 2006. 47(3): p. 432‐436.
Kautsky, H., et al., Neue versuche zur kohlensäureassimilation. Naturwissenschaften, 1931.
19(48): p. 964‐964.
Kautsky, H., et al., Chlorophyllfluoreszenz und koh‐lensäureassimilation x. die
chlorophyllfluoreszenz von ulva lactuca und ihre abhängigkeit von temperatur und lichtinten‐
sität. Biochem. Z, 1943. 315: p. 156‐175.
Kautsky, H., et al., Die fluoreszenzkurve und die photochemie der pflanze. Biochemistry, 1960.
332: p. 277‐292.
Pfündel, E., Estimating the contribution of photosystem I to total leaf chlorophyll
fluorescence. Photosynthesis Research, 1998. 56(2): p. 185‐195.
Stirbet, A., Chlorophyll a fluorescence induction: a personal perspective of the thermal phase,
the J–I–P rise. Photosynthesis Research, 2012. 113(1‐3): p. 15‐61.
Schansker, G., et al., Evidence for a fluorescence yield change driven by a light‐induced
conformational change within photosystem II during the fast chlorophyll a fluorescence rise.
Biochimica et Biophysica Acta (BBA) ‐ Bioenergetics, 2011. 1807(9): p. 1032‐1043.
Strasser, R.J., et al., Analysis of the chlorophyll a fluorescence transient. Advances in
Photosynthesis and Respiration. Vol. 19. 2004: Springer.
Ion transport in chloroplasts with role in regulation of photosynthesis
89.
90.
91.
92.
93.
94.
95.
96.
97.
98.
99.
100. 101. 102. 103. 104. 105. 106. 107. 108. Schreiber, U., et al., Continuous recording of photochemical and non‐photochemical
chlorophyll fluorescence quenching with a new type of modulation fluorometer.
Photosynthesis Research, 1986. 10(1‐2): p. 51‐62.
Maxwell, K., et al., Chlorophyll fluorescence—a practical guide. Journal of Experimental
Botany, 2000. 51(345): p. 659‐668.
Rascher, U., et al., Evaluation of instant light‐response curves of chlorophyll fluorescence
parameters obtained with a portable chlorophyll fluorometer on site in the field. Plant, cell &
environment, 2000. 23(12): p. 1397‐1405.
Ehleringer, J., Leaf absorptances of mohave and sonoran desert plants. Oecologia, 1981.
49(3): p. 366‐370.
Baker, N.R., Chlorophyll fluorescence: a probe of photosynthesis in vivo. Annual Review of
Plant Biology, 2008. 59: p. 89‐113.
Ruban, A.V., et al., The photoprotective molecular switch in the photosystem II antenna.
Biochimica et Biophysica Acta (BBA)‐Bioenergetics, 2012. 1817(1): p. 167‐181.
Nilkens, M., et al., Identification of a slowly inducible zeaxanthin‐dependent component of
non‐photochemical quenching of chlorophyll fluorescence generated under steady‐state
conditions in Arabidopsis. Biochimica et Biophysica Acta (BBA)‐Bioenergetics, 2010. 1797(4):
p. 466‐475.
Horton, P., et al., Allosteric regulation of the light‐harvesting system of photosystem II.
Philosophical Transactions of the Royal Society B: Biological Sciences, 2000. 355(1402): p.
1361‐1370.
Cazzaniga, S., et al., Interaction between avoidance of photon absorption, excess energy
dissipation and zeaxanthin synthesis against photooxidative stress in Arabidopsis. The Plant
Journal, 2013. 76(4): p. 568‐579.
Jensen, P.E., et al., The PSI‐K subunit of photosystem I is involved in the interaction between
light‐harvesting complex I and the photosystem I reaction center core. Journal of Biological
Chemistry, 2000. 275(32): p. 24701‐24708.
Tyystjärvi, E., Photoinhibition of photosystem II. International review of cell and molecular
biology, 2013. 300: p. 243‐303.
Duysens, L., Reversible changes in the absorption spectrum of chlorella upon irradiation. Science, 1954. 120(3113): p. 353‐354. Junge, W., et al., On the ion transport system of photosynthesis—Investigations on a molecular level—. Zeitschrift für Naturforschung B, 1968. 23(2): p. 244‐254. Schreiber, U., et al., New accessory for the Dual‐PAM‐100: The P515/535 module and examples of its application. PAM Application Notes (2008), 2008. 1: p. 1‐10. Bailleul, B., et al., Electrochromism: a useful probe to study algal photosynthesis. Photosynthesis Research, 2010. 106(1‐2): p. 179‐189. Cruz, J.A., et al., Contribution of electric field (Δψ) to steady‐state transthylakoid proton motive force (pmf) in vitro and in vivo. Control of pmf parsing into Δψ and ΔpH by ionic strength. Biochemistry, 2001. 40(5): p. 1226‐1237. Takizawa, K., et al., Depletion of stromal Pi induces high ‘energy‐dependent’antenna exciton quenching (qE) by decreasing proton conductivity at CFO‐CF1 ATP synthase. Plant, cell & environment, 2008. 31(2): p. 235‐243. Johnson, M.P., et al., Rethinking the existence of a steady‐state Δψ component of the proton motive force across plant thylakoid membranes. Photosynthesis Research, 2014. 119(1‐2): p. 233‐242. Vredenberg, W., et al., On the steady‐state electrical potential difference across the thylakoid membranes of chloroplasts in illuminated plant cells. Biochimica et Biophysica Acta (BBA)‐
Bioenergetics, 1975. 387(3): p. 580‐587. Enz, C., et al., Ion channels in the thylakoid membrane (a patch‐clamp study). Biochimica et Biophysica Acta (BBA)‐Bioenergetics, 1993. 1143(1): p. 67‐76. 51
Andrei Herdean
109. 110. 111. 112. 113. 114. 115. 116. 117. 118. 119. 120. 121. 122. 123. 124. 125. 126. 127. 128. 129. 130. 52
Schonknecht, G., et al., A voltage‐dependent chloride channel in the photosynthetic membrane of a higher plant. Nature, 1988. 336(6199): p. 589‐592. Avenson, T.J., et al., Regulating the proton budget of higher plant photosynthesis. Proceedings of the National Academy of Sciences, 2005. 102(27): p. 9709‐9713. Kramer, D.M., et al., Balancing the central roles of the thylakoid proton gradient. Trends in plant science, 2003. 8(1): p. 27‐32. Hochmal, A.K., et al., Calcium‐dependent regulation of photosynthesis. Biochimica et Biophysica Acta (BBA)‐Bioenergetics, 2015. Spetea, C., et al., Multiple evidence for nucleotide metabolism in the chloroplast thylakoid lumen. Proceedings of the National Academy of Sciences, 2004. 101(5): p. 1409‐1414. Spetea, C., et al., Evidence for nucleotide‐dependent processes in the thylakoid lumen of plant chloroplasts–an update. FEBS letters, 2012. 586(18): p. 2946‐2954. Thuswaldner, S., et al., Identification, expression, and functional analyses of a thylakoid ATP/ADP carrier from Arabidopsis. Journal of Biological Chemistry, 2007. 282(12): p. 8848‐
8859. Roth, C., et al., Characterization of a protein of the plastid inner envelope having homology to animal inorganic phosphate, chloride and organic‐anion transporters. Planta, 2004. 218(3): p. 406‐416. Schneider, A., et al., An Arabidopsis thaliana knock‐out mutant of the chloroplast triose phosphate/phosphate translocator is severely compromised only when starch synthesis, but not starch mobilisation is abolished. The Plant Journal, 2002. 32(5): p. 685‐699. Shutova, T., et al., Is the manganese stabilizing 33 kDa protein of photosystem II attaining a ‘natively unfolded’or ‘molten globule’structure in solution? FEBS letters, 2000. 467(2): p. 137‐
140. Kirchhoff, H., Architectural switches in plant thylakoid membranes. Photosynth Res, 2013. 116(2‐3): p. 481‐7. Yocum, C.F., The calcium and chloride requirements of the O2 evolving complex. Coordination Chemistry Reviews, 2008. 252(3): p. 296‐305. Olesen, K., et al., The function of the chloride ion in photosynthetic oxygen evolution. Biochemistry, 2003. 42(7): p. 2025‐2035. Ono, T.‐A., et al., Mn‐preserving extraction of 33‐, 24‐ and 16‐kDa proteins from O2‐evolving PS II particles by divalent salt‐washing. FEBS letters, 1983. 164(2): p. 255‐260. Redinbo, M., et al., Plastocyanin: structural and functional analysis. Journal of Bioenergetics and Biomembranes, 1994. 26(1): p. 49‐66. Nagy, G., et al., Chloroplast remodeling during state transitions in Chlamydomonas reinhardtii as revealed by noninvasive techniques in vivo. Proceedings of the National Academy of Sciences, 2014. 111(13): p. 5042‐5047. Goss, R., et al., The importance of grana stacking for xanthophyll cycle‐dependent NPQ in the thylakoid membranes of higher plants. Physiologia plantarum, 2007. 131(3): p. 496‐507. Barber, J., An explanation for the relationship between salt‐induced thylakoid stacking and the chlorophyll fluorescence changes associated with changes in spillover of energy from photosystem II to photosystem I. FEBS letters, 1980. 118(1): p. 1‐10. Barber, J., et al., Electrical diffuse layers and their influence on photosynthetic processes. FEBS letters, 1977. 74(2): p. 174‐181. Barber, J., Membrane surface charges and potentials in relation to photosynthesis. Biochimica et Biophysica Acta (BBA) ‐ Reviews on Bioenergetics, 1980. 594(4): p. 253‐308. Rumak, I., et al., 3‐D modelling of chloroplast structure under (Mg2+) magnesium ion treatment. Relationship between thylakoid membrane arrangement and stacking. Biochimica et Biophysica Acta (BBA) ‐ Bioenergetics, 2010. 1797(10): p. 1736‐1748. Barber, J., et al., A mechanism for controlling the stacking and unstacking of chloroplast thylakoid membranes. FEBS letters, 1979. 105(1): p. 5‐10. Ion transport in chloroplasts with role in regulation of photosynthesis
131. 132. 133. 134. 135. 136. 137. 138. 139. 140. Jennings, R.C., et al., Studies on cation‐induced thylakoid membrane stacking, fluorescence yield, and photochemical efficiency. Plant physiology, 1978. 62(6): p. 879‐884. Hind, G., et al., Light‐dependent redistribution of ions in suspensions of chloroplast thylakoid membranes. Proceedings of the National Academy of Sciences, 1974. 71(4): p. 1484‐1488. Ettinger, W.F., et al., Identification of a Ca2+/H+ antiport in the plant chloroplast thylakoid membrane. Plant physiology, 1999. 119(4): p. 1379‐1386. Tapken, W., et al., The Clp protease system is required for copper ion‐dependent turnover of the PAA2/HMA8 copper transporter in chloroplasts. New Phytologist, 2015. 205(2): p. 511‐
517. Alscher, R.G., et al., Role of superoxide dismutases (SODs) in controlling oxidative stress in plants. Journal of Experimental Botany, 2002. 53(372): p. 1331‐1341. Chen, M., et al., Mutations in the Arabidopsis VAR2 locus cause leaf variegation due to the loss of a chloroplast FtsH protease. The Plant Journal, 2000. 22(4): p. 303‐313. Ostersetzer, O., et al., Light‐stimulated degradation of an unassembled Rieske FeS protein by a thylakoid‐bound protease: the possible role of the FtsH protease. The Plant Cell, 1997. 9(6): p. 957‐65.
Dumas, R., et al., Evidence for two catalytically different magnesium‐binding sites in acetohydroxy acid isomeroreductase by site‐directed mutagenesis. Biochemistry, 1995. 34(18): p. 6026‐6036. Malyan, A., et al., Mg2+ ‐dependent inactivation / H+ ‐dependent activation equilibrium of chloroplast F1‐ATPase. Photosynthesis Research, 1998. 57(3): p. 297‐303. Reinhardt, K., et al., Clouds and cloud immersion alter photosynthetic light quality in a temperate mountain cloud forest. Botany, 2010. 88(5): p. 462‐470. 53